Please help!
Recently our SYBR Green qPCR reaction using our housekeeping gene, RPLP0, and all the same conditions stopped working one day to the next.
We isolate RNA from tissue culture human cells via Trizol-chloroform extraction, treat with Invitrogen DNase I, and use 1ug of DNase treated RNA in a Superscript IV RT reaction. We then dilute cDNA 1:4 for use with the Applied Biosystems Power SYBR Green mix.
Recently, only the first undiluted point of our standard curve amplifies - every 1:10 dilution after that only comes up with a very flat amplification curve. We've tried a different lot of SYBR Green, ordered new primers (and run them out on an agarose gel to make sure they weren't degraded), different primers, new water, made new cDNA, not DNase treating... I'm at wit's end! We have used these exact samples before and gotten a beautiful standard curve using the same primers, master mix, and run parameters.
I've taken the products from a qPCR run and run them out on an agarose gel, and nothing seems to be amplifying in the 1:10-1:1000 dilutions. What could be inhibiting the reactions in only those dilutions?
Real-time PCR is a specialized technique that delivers far more information about your DNA or RNA than end-point PCR.
Essentially, real-time PCR is a way to visualize the amplification of specific DNA fragments as it is happening (in real time) and allows for the ability to quantify exactly how much DNA (or RNA) was in the original sample.
Fluorescent dyes (either SYBR Green or dye labeled DNA oligos) are mixed in the amplification reaction, fluoresce when they bind to DNA, and are measured by the machine during replication.
The amount of fluorescent signal measured directly correlates to the amount of product in the tube at that moment in time. Because of the sensitivity of fluorescent dyes, concentrations as low as picogram quantities of DNA (or as low as a single cell) can be accurately detected.
While it has become second nature in some labs, the technique does require a certain amount of technical finesse to get consistent real-time PCR data every time. Because the cost of real-time PCR kits is so much higher than standard PCR, getting every experiment right is critical.
The main bottlenecks people encounter when getting started with real-time PCR are contamination issues or inconsistency between replicates. Here are some simple but important pointers to help make sure you prevent the cause of many grey hairs and frustration at lab meetings.
1. Always Mix the Reagents Well Before Use
The reagents contain dyes, nucleotides, and enzymes that may have settled while sitting in the freezer or refrigerator. Give the master mix a good mix before your start aliquoting into your plate or tubes to avoid uneven distribution of reagents between samples.
2. Store Primers in a Buffer to Protect their Stability
When your primers arrive, avoid resuspending the master stock in water. The pH of water can be low (especially if it is DEPC treated) and this will be damaging to DNA over time.
Use a buffered solution at neutral pH to protect from acid hydrolysis. EDTA (1 mM) in the master stock is also a good idea to protect against DNases and when you dilute the primers for working stocks, the EDTA will be sufficiently dilute that it will not interfere with Taq activity.
3. Aliquot the Primers
Once you have a master stock (usually 100-200 mM), you will want to make some working stocks so that you are not continually freeze/thawing your primary source. Prepare 10-20 mM working stocks in neutral pH buffer and prepare aliquots that allow you to freeze/thaw the working stock 3-5 times at the most.
Continual freeze/thawing of the primers can cause some break down and this will lead to drift in results such as worsening PCR efficiency and sensitivity. Preparing aliquots will also help avoid contamination problems.
If you accidentally contaminate one of the tubes of primer, you can throw it away and take a fresh one without worrying about contaminating the main stock.
4. Use Pipettors Accurately for Low Volumes
If you require absolute accuracy in quantification and want spot on standard curves, use a pipettor calibrated for low volume pipetting (such as a P2 or P10).
This will ensure reproducibility between replicates and make sure that when you are measuring efficiency based on the standard curve, that you are truly measuring efficiency of the reaction and not your pipetting skill. Read more to learn more about pipetting accuracy.
5. Perform a Standard Curve for Every New Primer Pair
Don’t assume that every set of primers ordered is going to work as well as the last. PCR efficiency can be impacted by a number of factors. The best practice is to run a 5 point standard curve with 10 fold dilutions for every new primer pair and make sure you can get at least 90% PCR efficiency with control DNA.
7. Follow the Three Room Rule
One of the biggest causes of contamination is from using the same pipettors for extraction or handling PCR products post-run for reaction set up. Even if aerosol resistant tips are used all the time, this is a big no-no. Buy a complete set of pipettors that are used for PCR set up and nothing else.
In addition to new pipettors, you will want to keep them in a different location, and preferably a different room than the room used for extractions. The ideal set up is to have three rooms; one for RNA or DNA extractions, one for reaction set up (and using a hood with a UV lamp to pre-treat the pipettors and plastics between users), and one for the real-time cycler.
This is the most assured way to make sure you never have amplification in your negative controls.
8. Double Check the Cycling Conditions
This is important if you are using a shared instrument. Even if you have your own template file set up, before hitting start, make sure the machine has the correct run cycle for your experiment. Someone may have used your template and made changes to the annealing temperature or the hot start activation time without your knowledge.
Some instruments default back to standard settings and if you are using an instrument for the first time, you may find that your settings didn’t save. It never hurts to double check the run settings.
9. Dilute the Template (Less is More)
Depending on the gene of interest, you might actually be starting with too much template. Real-time PCR is sensitive enough that sometimes less template gives a more accurate measurement.
You will want samples to cross the threshold between cycles 20-30. Samples that cross the threshold below cycle 15 will fall into most instruments default baseline setting and this will cause a subtraction of fluorescence from the rest of the data.
This can be remedied by adjusting the baseline setting, but if you are unfamiliar with your instrument, it may require a call to technical service to figure it out. Also, if there were any inhibitors in the sample from the purification step (guanidine salts or ethanol, for example) diluting the sample will eliminate their impact on the results and give you an accurate quantitation of the sample.
The best approach for a new sample is to perform a standard curve- even just a 3 point dilution series- to see what concentration will give you a Ct that falls in your standard curve and is most accurate.
10. Make Dilutions Fresh
Nucleic acids stick to plastic so if you make a dilution series and want to store it for future runs, you will need to protect the samples from absorbing to the tubes walls and becoming diluted out over time.
This can be done by using a carrier nucleic acid, such as tRNA, or by using specially treated plasticware that does not bind nucleic acids. Several manufacturers offer low retention tubes (Axygen is one) or silicon treated tubes to help prevent this occurrence.
If you do store dilutions in non-treated tubes, you may want to re-quantify the most concentrated dilutions on a Nanodrop before using to make sure they still match the expected concentration.
Some of these tips may seem like common sense- and they are- to people who have been doing this for a long time. But for many people just starting to use this technology, a lot of time and money can be saved with these simple steps that can make a big impact on results.
There are a lot of resources for real-time PCR help as well, including a very active Yahoo List group and the BioTechniques® Molecular Biology Forums. Fortunately, there are many experts who enjoy helping others master the art of real-time PCR.
And if any experts out there reading this want to list some common mistakes you see in your labs or best practices tips, please let us know in the comments field.
Real-time PCR is a specialized technique that delivers far more information about your DNA or RNA than end-point PCR.
Essentially, real-time PCR is a way to visualize the amplification of specific DNA fragments as it is happening (in real time) and allows for the ability to quantify exactly how much DNA (or RNA) was in the original sample.
Fluorescent dyes (either SYBR Green or dye labeled DNA oligos) are mixed in the amplification reaction, fluoresce when they bind to DNA, and are measured by the machine during replication.
The amount of fluorescent signal measured directly correlates to the amount of product in the tube at that moment in time. Because of the sensitivity of fluorescent dyes, concentrations as low as picogram quantities of DNA (or as low as a single cell) can be accurately detected.
While it has become second nature in some labs, the technique does require a certain amount of technical finesse to get consistent real-time PCR data every time. Because the cost of real-time PCR kits is so much higher than standard PCR, getting every experiment right is critical.
The main bottlenecks people encounter when getting started with real-time PCR are contamination issues or inconsistency between replicates. Here are some simple but important pointers to help make sure you prevent the cause of many grey hairs and frustration at lab meetings.
1. Always Mix the Reagents Well Before Use
The reagents contain dyes, nucleotides, and enzymes that may have settled while sitting in the freezer or refrigerator. Give the master mix a good mix before your start aliquoting into your plate or tubes to avoid uneven distribution of reagents between samples.
2. Store Primers in a Buffer to Protect their Stability
When your primers arrive, avoid resuspending the master stock in water. The pH of water can be low (especially if it is DEPC treated) and this will be damaging to DNA over time.
Use a buffered solution at neutral pH to protect from acid hydrolysis. EDTA (1 mM) in the master stock is also a good idea to protect against DNases and when you dilute the primers for working stocks, the EDTA will be sufficiently dilute that it will not interfere with Taq activity.
3. Aliquot the Primers
Once you have a master stock (usually 100-200 mM), you will want to make some working stocks so that you are not continually freeze/thawing your primary source. Prepare 10-20 mM working stocks in neutral pH buffer and prepare aliquots that allow you to freeze/thaw the working stock 3-5 times at the most.
Continual freeze/thawing of the primers can cause some break down and this will lead to drift in results such as worsening PCR efficiency and sensitivity. Preparing aliquots will also help avoid contamination problems.
If you accidentally contaminate one of the tubes of primer, you can throw it away and take a fresh one without worrying about contaminating the main stock.
4. Use Pipettors Accurately for Low Volumes
If you require absolute accuracy in quantification and want spot on standard curves, use a pipettor calibrated for low volume pipetting (such as a P2 or P10).
This will ensure reproducibility between replicates and make sure that when you are measuring efficiency based on the standard curve, that you are truly measuring efficiency of the reaction and not your pipetting skill. Read more to learn more about pipetting accuracy.
5. Perform a Standard Curve for Every New Primer Pair
Don’t assume that every set of primers ordered is going to work as well as the last. PCR efficiency can be impacted by a number of factors. The best practice is to run a 5 point standard curve with 10 fold dilutions for every new primer pair and make sure you can get at least 90% PCR efficiency with control DNA.
7. Follow the Three Room Rule
One of the biggest causes of contamination is from using the same pipettors for extraction or handling PCR products post-run for reaction set up. Even if aerosol resistant tips are used all the time, this is a big no-no. Buy a complete set of pipettors that are used for PCR set up and nothing else.
In addition to new pipettors, you will want to keep them in a different location, and preferably a different room than the room used for extractions. The ideal set up is to have three rooms; one for RNA or DNA extractions, one for reaction set up (and using a hood with a UV lamp to pre-treat the pipettors and plastics between users), and one for the real-time cycler.
This is the most assured way to make sure you never have amplification in your negative controls.
8. Double Check the Cycling Conditions
This is important if you are using a shared instrument. Even if you have your own template file set up, before hitting start, make sure the machine has the correct run cycle for your experiment. Someone may have used your template and made changes to the annealing temperature or the hot start activation time without your knowledge.
Some instruments default back to standard settings and if you are using an instrument for the first time, you may find that your settings didn’t save. It never hurts to double check the run settings.
9. Dilute the Template (Less is More)
Depending on the gene of interest, you might actually be starting with too much template. Real-time PCR is sensitive enough that sometimes less template gives a more accurate measurement.
You will want samples to cross the threshold between cycles 20-30. Samples that cross the threshold below cycle 15 will fall into most instruments default baseline setting and this will cause a subtraction of fluorescence from the rest of the data.
This can be remedied by adjusting the baseline setting, but if you are unfamiliar with your instrument, it may require a call to technical service to figure it out. Also, if there were any inhibitors in the sample from the purification step (guanidine salts or ethanol, for example) diluting the sample will eliminate their impact on the results and give you an accurate quantitation of the sample.
The best approach for a new sample is to perform a standard curve- even just a 3 point dilution series- to see what concentration will give you a Ct that falls in your standard curve and is most accurate.
10. Make Dilutions Fresh
Nucleic acids stick to plastic so if you make a dilution series and want to store it for future runs, you will need to protect the samples from absorbing to the tubes walls and becoming diluted out over time.
This can be done by using a carrier nucleic acid, such as tRNA, or by using specially treated plasticware that does not bind nucleic acids. Several manufacturers offer low retention tubes (Axygen is one) or silicon treated tubes to help prevent this occurrence.
If you do store dilutions in non-treated tubes, you may want to re-quantify the most concentrated dilutions on a Nanodrop before using to make sure they still match the expected concentration.
Some of these tips may seem like common sense- and they are- to people who have been doing this for a long time. But for many people just starting to use this technology, a lot of time and money can be saved with these simple steps that can make a big impact on results.
There are a lot of resources for real-time PCR help as well, including a very active Yahoo List group and the BioTechniques® Molecular Biology Forums. Fortunately, there are many experts who enjoy helping others master the art of real-time PCR.
And if any experts out there reading this want to list some common mistakes you see in your labs or best practices tips, please let us know in the comments field.
Justine, did you run these reactions only on a single instrument? Or did you try to run it on a different instrument?
Since it seems like you're trying to check every variable here, that may be one other variable to look at. Sometimes instruments can have a failed heating or cooling element in the block that can affect amplification.
Have you also run a positive control? If you haven't, you may just want to purchase a positive control, and run this at different dilutions to see whether you are getting good amplification from something that you know should work.
If it worked beautifully before, but is now giving issues with all the same reagents and machine my best guess is either: your qPCR reagents have been frozen and thawed too many times so they don't work. OR someone changed the program in your thermocycler.
Hi Justine, your plot shows very consistent pipetting and you have eliminated many possibilities already, which is very good! Have you tried doing a standard curve with genomic DNA instead using the exact same reagents, parameters and dilutions? Also, this other RG question from 4 years ago might aid you on your quest:
Good luck!
https://www.researchgate.net/post/Anyone_seen_qPCR_curves_like_this
I agree with the equipment failure possibility. Run a basic pct in the machine to test the wells
I agree with the equipment failure possibility. Run a basic pct in the machine to test the wells performance and uniformity. Use a different machine to set supplies.
So you have tried dilution curve method and did not observe anything on the agarose gel for the dilutions. Try doing a simple PCR reaction with same cDNA input (with the serial dilutions of the undiluted sample) with normal PCR reagents and see if it works. If it works, this would hint towards possible problems in the SYBR reaction reagents, qPCR run (protocol, although your undiluted sample gives nice amplification). Have you checked the Ct values in the export file???
Dear Justine, I will suggest you to recheck your cDNA concentration stock and prepare a fresh cDNA dilution, because you said that your first undiluted point of standard curve amplifies and rest one having the problem. It might be a dilution error. Even though it’s not working you can go with Debatosh Das idea doing a simple PCR reaction with same cDNA input to rule out the SYBR green reagents problem.
Hi Justine,
I think if you alredy tried the primers, the qPCR protocol, and the master mix, it is likely that you have a contaminated cDNA (DNase?). I would clean everything with RNase/DNase Away, and start from scratch. Including your cDNA or plasmids. And I think Lalan is right, check the dilutions again.
Good luck!
Hi Justine,
Have you found what was the problem?, because I have the same trouble and I don't know what what else to do
Tamara D Keenum and Isabel Cervantes, were you able to overcome the problem with your qPCR? Would you mind sharing how you got it solved? Thank you.
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